Polymer-Degrading Enzymes of Pseudomonas chloroaphis PA23 Display Broad Substrate Preferences

Although many bacterial lipases and PHA depolymerases have been identified, cloned, and characterized, there is very little information on the potential application of lipases and PHA depolymerases, especially intracellular enzymes, for the degradation of polyester polymers/plastics. We identified genes encoding an intracellular lipase (LIP3), an extracellular lipase (LIP4), and an intracellular PHA depolymerase (PhaZ) in the genome of the bacterium Pseudomonas chlororaphis PA23. We cloned these genes into Escherichia coli and then expressed, purified, and characterized the biochemistry and substrate preferences of the enzymes they encode. Our data suggest that the LIP3, LIP4, and PhaZ enzymes differ significantly in their biochemical and biophysical properties, structural-folding characteristics, and the absence or presence of a lid domain. Despite their different properties, the enzymes exhibited broad substrate specificity and were able to hydrolyze both short- and medium-chain length polyhydroxyalkanoates (PHAs), para-nitrophenyl (pNP) alkanoates, and polylactic acid (PLA). Gel Permeation Chromatography (GPC) analyses of the polymers treated with LIP3, LIP4, and PhaZ revealed significant degradation of both the biodegradable as well as the synthetic polymers poly(ε-caprolactone) (PCL) and polyethylene succinate (PES).


Introduction
The Pseudomonadaceae are a family of Gram-negative aerobic chemoheterotrophs that possess diverse enzyme systems and are capable of many biochemical transformations. Members of the metabolically diverse genus Pseudomonas have gained particular interest due to their capabilities to degrade and metabolize synthetic plastics [1]. Pseudomonas chlororaphis PA23 is a known biocontrol agent that exhibits plant growth promotion potential and inhibits the growth of plant fungal pathogens by secreting an arsenal of antibiotics and degradative enzymes [2,3].
Lipase enzymes are known to catalyze the hydrolysis of triacylglycerides (TAGs) to free fatty acids and glycerol. In addition, some lipases are able to synthesize new products by esterification, transesterification, and aminolysis [6,7]. Lipases act by "interfacial activation" where the aqueous phase containing the enzyme meets the insoluble, hydrophobic surface of the substrate [6]. This ability to function with enhanced activity at the nonpolar hydrophobic-aqueous hydrophilic interface is a property that separates lipases from other esterases [7].

Cloning, Expression and Purification of LIP3, LIP4 and PhaZ
The complete genome sequence of Pseudomonas chlororaphis PA23 is available in NCBI (https://www.ncbi.nlm.nih.gov (accessed on 1 July 2021)) and Integrated Microbial Genome (IMG) (https://img.jgi.doe.gov (accessed on 1 July 2021)) (Genome Acc. No. CP008696). Careful examination revealed seven genes encoding esterase/lipase enzymes in the P. chlororaphis PA23 genome ( Table 1). Three of these encode intracellular lipases (EY04_08410, EY04_09635, and EY04_02420), while the other three contain a signal peptide indicative of possible extracellular activity (EY04_21540, EY04_17885, and EY04_32435). Curiously, only one encodes an intracellular PHA depolymerase (EY04_01535). In our initial experiments with P. chlororaphis PA23 and Acinetobacter lwoffii, we found that hydrolysis of the ester bonds of pNP-alkanoate substrates and the ester bonds of PHA polymers with different subunit compositions by the extracellular esterases/lipases was detectable in the growth medium [24]. In the present study, two genes encoding an intracellular and extracellular lipaseand an intracellular PHA depolymerase were cloned and expressed in E. coli BL21(DE3) to compare the properties and investigate their biodegradability compared to different biodegradable polymers. We used the pET28a(+) vector to clone the lip3, lip4, and phaZ genes using Escherichia coli BL21(DE3) as the expression host. The main expression/fermentation parameter was that optimal expression was achieved after 5 h with 0.6 mmol L −1 IPTG at 37 • C ( Figure 1A). After fermentation with expression, the cells were harvested and the recombinant proteins were purified to homogeneity. We roughly estimated the molecular masses by SDS-PAGE analysis of the purified enzymes and found molecular masses of 32 to 38 kDa for LIP3, LIP4, and PhaZ. These masses were consistent with the predicted molecular masses from their amino acid sequences (https://web.expasy.org/protparam (accessed on 15 September 2021)) ( Figure 1A,B).
Purified enzymes were then applied to agar plates with colloidal overlays of different polymers to assess their polymer-degrading abilities. Control treatments consisted of soluble crude extracts of uninduced recombinant E. coli BL21 cells containing lip3-pET28a(+)/lip4-pET28a(+)/phaZ-pET28a(+). Enzymatic hydrolysis of the polymers was verified by observing the formation of fluorescent "zones of clearance" around the wells after 48 h incubation at 30 • C under UV irradiation at 350 nm ( Figure 2). These experiments clearly demonstrated that the purified enzymes were able to hydrolyze PHA polymers.  PHA agar plate assay for testing the depolymerase/esterase activity of LIP3, LIP4, and PhaZ. The clearance zones around the wells to which 20 µ g of purified LIP3, LIP4, and PhaZ enzymes were added show hydrolysis of mcl-PHA in rhodamine B-PHO agar plates after UV irradiation. As a control, a crude extract of the noninduced expression host E. coli BL21(DE3) was used.

Sequence Analyses and Homology Modeling of LIP3, LIP4, and PhaZ
The lip3, lip4, and phaZ genes encode enzymes with predicted molecular masses of 32.5 kDa, 34.0 kDa, and 31.7 kDa, respectively. BLAST analyses of the deduced amino acid sequences (BLASTp) confirmed that the LIP3 and LIP4 enzymes belong to the lipase superfamily, whereas PhaZ belongs to the depolymerase superfamily, and all three have an α/β-hydrolase fold. Alignment of the amino acid sequences of LIP3 and LIP4 revealed very low amino acid sequence identity (12.29%), LIP3 and PhaZ had an amino acid sequence identity of 14.44%, and LIP4 and PhaZ showed an amino acid sequence identify  PHA agar plate assay for testing the depolymerase/esterase activity of LIP3, LIP4, and PhaZ. The clearance zones around the wells to which 20 µ g of purified LIP3, LIP4, and PhaZ enzymes were added show hydrolysis of mcl-PHA in rhodamine B-PHO agar plates after UV irradiation. As a control, a crude extract of the noninduced expression host E. coli BL21(DE3) was used.

Sequence Analyses and Homology Modeling of LIP3, LIP4, and PhaZ
The lip3, lip4, and phaZ genes encode enzymes with predicted molecular masses of 32.5 kDa, 34.0 kDa, and 31.7 kDa, respectively. BLAST analyses of the deduced amino acid sequences (BLASTp) confirmed that the LIP3 and LIP4 enzymes belong to the lipase superfamily, whereas PhaZ belongs to the depolymerase superfamily, and all three have an α/β-hydrolase fold. Alignment of the amino acid sequences of LIP3 and LIP4 revealed very low amino acid sequence identity (12.29%), LIP3 and PhaZ had an amino acid sequence identity of 14.44%, and LIP4 and PhaZ showed an amino acid sequence identify Figure 2. PHA agar plate assay for testing the depolymerase/esterase activity of LIP3, LIP4, and PhaZ. The clearance zones around the wells to which 20 µg of purified LIP3, LIP4, and PhaZ enzymes were added show hydrolysis of mcl-PHA in rhodamine B-PHO agar plates after UV irradiation. As a control, a crude extract of the noninduced expression host E. coli BL21(DE3) was used.

Sequence Analyses and Homology Modeling of LIP3, LIP4, and PhaZ
The lip3, lip4, and phaZ genes encode enzymes with predicted molecular masses of 32.5 kDa, 34.0 kDa, and 31.7 kDa, respectively. BLAST analyses of the deduced amino acid sequences (BLASTp) confirmed that the LIP3 and LIP4 enzymes belong to the lipase superfamily, whereas PhaZ belongs to the depolymerase superfamily, and all three have an α/β-hydrolase fold. Alignment of the amino acid sequences of LIP3 and LIP4 revealed very low amino acid sequence identity (12.29%), LIP3 and PhaZ had an amino acid sequence identity of 14.44%, and LIP4 and PhaZ showed an amino acid sequence identify of 9.86%. However, all three enzymes have conserved amino acid sequence motifs corresponding to the lipase box (GX 1 SX 2 G) characteristic of the lipase/esterase family and the putative oxyanion hole residues (GMLG-in LIP3; HGGGW-in LIP4, and PFV-in PhaZ) in the catalytic domain (Figures 3-5). The x 1 sites in the lipase boxes of LIP3, LIP4, and PhaZ are occupied by histidine (H82), asparagine (N179), and valine (V101), while the x 2 sites contain glutamine (Q84), valine (V181), and tryptophan (W103), respectively. The enzymes possess a catalytic triad (serine-histidine-aspartic acid) characteristic of other lipases. The catalytic triad LIP3 has Ser 83 -His 263 -Asp 241 (Figure 3), the catalytic triad of LIP4 has Ser 180 -His 307 -Asp 277 (Figure 4), while PhaZ has Ser 102 -His 248 -Asp 221 in its active site ( Figure 5).  (Figure 3), the catalytic triad of LIP4 has Ser 180 -His 307 -Asp 277 (Figure 4), while PhaZ has Ser 102 -His 248 -Asp 221 in its active site ( Figure 5). and Pseudomonas aeruginosa (1EX9). Brown box: amino acids of the lipase box consensus sequence; Blue box: amino acid residues of the oxyanion hole; green box: amino acid residues of the closing lid domain; blue stars: show the positions of the amino acids of the catalytic triad (serine, glutamic acid, and histidine); secondary structure α-helix and β-sheet regions are shown alone at the top of the alignment. White letters with red highlights represent amino acid residues that are conserved in all the enzyme sequences and red letters in blue frame indicate conserved regions with identical amino acid residues (residues with similar properties) in one or two of the enzyme sequences.  Brown box: amino acids of the consensus sequence of the lipase box; blue box: amino acid residues of the oxyanion hole; green box: amino acid residues of the closing lid domain; blue asterisks: the positions of the amino acids of the catalytic triad (serine, glutamic acid, and histidine) in the sequence; the secondary structure segments of the α-helix and the β-sheet are shown alone at the top of the alignment. White letters with red highlights represent amino acid residues that are conserved in all the enzyme sequences and red letters in blue frame indicate conserved regions with identical amino acid residues (residues with similar properties) in one or two of the enzyme sequences.  PSI-BLAST search for LIP3, LIP4, and PhaZ in the protein data bank revealed several bacterial lipases and PHA depolymerases as close homologues. Lipase from Proteus mirabilis (4GW3) was chosen as the template for LIP3 (43% amino acid identity and 95% similarity), lipase CinB from Enterobacter asburiae, which is an acetyl esterase (6KMO), was chosen for LIP4 (67% amino acid identity and 98% similarity), and PHA depolymerase from Pseudomonas oleovorans was chosen for PhaZ (91% amino acid identity and 98% similarity).
Supplementary Figures S1A, S2A, and S3A show the homology model of LIP3, LIP4, and PhaZ, respectively, with the substrate 3-hydroxyoctanoate. The core domain of LIP3 consists of a five-parallel β-sheet, whereas LIP4 and PhaZ consists of an eight-parallel Figure 5. Amino acid sequence alignment of PhaZ with the serine hydrolase CCSP0084 from Cycloclasticus sp. P1 (4I3F). Brown box: amino acids of the Lipase box consensus sequence; blue box: amino acid residues of the oxyanion hole; blue asterisks indicate the positions of the amino acids of the catalytic triad (serine, glutamic acid, and histidine); the sequence parts corresponding with α-helix and β-sheet regions are shown alone at the top of the alignment. White letters with red highlights represent amino acid residues that are conserved in all the enzyme sequences and red letters in blue frame indicate conserved regions with identical amino acid residues (residues with similar properties) in one or two of the enzyme sequences. The grey asterisks above amino acid residues 67, 88, 126, and 192 of both sequences, 75 and 126 of the PhaZ sequence, 174 of the 4I3F/A sequence, indicate positions with amino acids with similar properties.
PSI-BLAST search for LIP3, LIP4, and PhaZ in the protein data bank revealed several bacterial lipases and PHA depolymerases as close homologues. Lipase from Proteus mirabilis (4GW3) was chosen as the template for LIP3 (43% amino acid identity and 95% similarity), lipase CinB from Enterobacter asburiae, which is an acetyl esterase (6KMO), was chosen for LIP4 (67% amino acid identity and 98% similarity), and PHA depolymerase from Pseudomonas oleovorans was chosen for PhaZ (91% amino acid identity and 98% similarity).
Supplementary Figures S1A, S2A and S3A show the homology model of LIP3, LIP4, and PhaZ, respectively, with the substrate 3-hydroxyoctanoate. The core domain of LIP3 consists of a five-parallel β-sheet, whereas LIP4 and PhaZ consists of an eight-parallel β-sheet (except for strand two, which is antiparallel).In each model, there are multiple αhelices on each side of the core β-sheet. Supplementary Figures S1B, S2B and S3B show the binding pocket of the substrate 3-hydroxyoctanoate dimer (residues 130-148 and 221-232 of LIP3, and 23-63 of LIP4), while the substrate binding pocket of PhaZ is exposed and lacks the closing lid domain.
To better illustrate the substrate binding pocket, the closing lid domain is not shown in Supplementary Figures S1B and S2B. It should also be noted that Supplementary Figures S1C, S2C and S3C present the surface of the lipases and PhaZ, clearly showing that the substrate binding pocket for LIP3 and LIP4 is buried by the lid domain. The binding energy for the substrate to LIP3, LIP4, and PhaZ determined in the docking experiment was calculated to be −6.2 kcal/mol, −5.8 kcal/mol, and −5.7 kcal/mol, respectively.A negative free energy of binding means that the substrates dock spontaneously.
β-sheet (except for strand two, which is antiparallel).In each model, there are multiple α-helices on each side of the core β-sheet. Supplementary Figures S1B, S2B,and S3B show the binding pocket of the substrate 3-hydroxyoctanoate dimer (residues 130-148 and 221-232 of LIP3, and 23-63 of LIP4), while the substrate binding pocket of PhaZ is exposed and lacks the closing lid domain.
To better illustrate the substrate binding pocket, the closing lid domain is not shown in Supplementary Figures S1B and S2B.It should also be noted that Supplementary Figure S1C, S2C and S3C present the surface of the lipases and PhaZ, clearly showing that the substrate binding pocket for LIP3 and LIP4 is buried by the lid domain. The binding energy for the substrate to LIP3, LIP4, and PhaZ determined in the docking experiment was calculated to be −6.2 kcal/mol, −5.8 kcal/mol, and −5.7 kcal/mol, respectively.A negative free energy of binding means that the substrates dock spontaneously.
LIP3, LIP4, and PhaZ were optimally active at 45 °C to 50 °C and retained more than 70% activity when the reaction was performed at 55 °C ( Figure 6B). In addition, thermal inactivation experiments showed that the enzymes retained nearly 70% activity for 24 h at 45 °C ( Figure 6C). LIP3 and PhaZ, on the other hand, were very stable and retained more than 80% of their activity for 24 h at 45 °C.LIP3 exhibited higher activation energy (Ea = 28.2 KJ mol −1 ) than LIP4 (26.4 KJ mol −1 ) and PhaZ (19.9 KJ mol −1 ) for the thermo-inactivation process within the temperature range of 25-50 °C. LIP3, LIP4, and PhaZ were optimally active at 45 • C to 50 • C and retained more than 70% activity when the reaction was performed at 55 • C ( Figure 6B). In addition, thermal inactivation experiments showed that the enzymes retained nearly 70% activity for 24 h at 45 • C ( Figure 6C). LIP3 and PhaZ, on the other hand, were very stable and retained more than 80% of their activity for 24 h at 45 • C. LIP3 exhibited higher activation energy (E a = 28.2 KJ mol −1 ) than LIP4 (26.4 KJ mol −1 ) and PhaZ (19.9 KJ mol −1 ) for the thermo-inactivation process within the temperature range of 25-50 • C.
When pNP-octanoate was used as a substrate, the purified enzymes were found to be very stable and retained almost 100% of their activities after four months of incubation at 4 • C and two months of incubation at 30 • C. The pNP-octanoate was also used as a substrate to determine the kinetics of each enzyme. The V max and K m for LIP3 were 625.0 µM mg −1 min −1 and 0.375 mmol L −1 , respectively; the V max and K m of LIP4 were 555.55µM mg −1 min −1 and 0.333 mmol L −1 , respectively; and the V max and K m of PhaZ were 909.09 µM mg −1 min −1 and 0.636 mmol L −1 , respectively.
When the effect of cations and other compounds was examined (Table 2), it was found that 1 mmol L −1 CaCl 2 , 10 mmol L −1 EDTA and other metal ions did not affect the enzyme's activity, indicating that the enzymes are not metalloenzymes and do not require metal ions (calcium, sodium, and magnesium) as cofactors for enzyme activity. PMSF and ionic and nonionic detergents (SDS and Tween 80) inhibited the activities of LIP3, LIP4, and PhaZ.

Substrate Specificity of LIP3, LIP4, and PhaZ
The lipase enzymes LIP3 and LIP4 as well as the PHA depolymerase PhaZ showed broad substrate specificity. In particular, the enzymes showed significant activity towards the ester bonds of pNP-alkanoates, which are the typical substrate for lipases ( Figure 7). The enzymes hydrolyzed pNP-alkanoates with shorter and longer side chains, and the esterase activity for pNP octanoate was highest for LIP3 and PhaZ, whereas LIP4 showed the highest esterase activity for pNP-acetate. The substrate specificity of LIP3 and PhaZ for medium-and long-chain pNP-alkanoates was relatively higher than for short chain pNP-alkanoates. LIP4, on the other hand, showed higher specificity (>70%) for short chain pNP-alkanoates (pNP-acetate and pNP-butyrate).
Turbidimetric assays demonstrated that purified LIP3, LIP4, and PhaZ hydrolyzed ester linkages of β-polyhydroxyalkanoates/PHA polymers prepared from valeric acid (PHBV), hexanoic acid (PHHx), octanoic acid (PHO), nonanoic acid (PHN), and decanoic acid (PHD) ( Table 3). The substrate preference for the LIP3 was PHHx > PHBV > PHO > PHN > PHD, the substrate preference for LIP4 was PHHx > PHO > PHN > PHD > PHBV, andthe substrate preference for PhaZ was PHO > PHN > PHD > PHHx > PHBV. Using PHO as substrate, LIP3 had a specific activity of 122.13 units PHA mg −1 protein, LIP4 had a specific activity of 102.6 units PHA mg −1 protein, and PhaZ had a specific activity of 650.13 units PHA mg −1 protein. The data reveal that the LIP3, LIP4, and PhaZ enzymes have a wide-range of substrate preferences, including scl-PHA polymers and mcl-PHA polymers, as well as C4 to C10 pNP-alkanoates. These enzymes also showed high activity towards PLA and were also able to hydrolyze synthetic polyester polymers, such as PCL and PES (Table 3). andthe substrate preference for PhaZ was PHO > PHN > PHD > PHHx > PHBV. Using PHO as substrate, LIP3 had a specific activity of 122.13 unitsPHA mg −1 protein, LIP4 had a specific activity of 102.6 unitsPHA mg −1 protein, and PhaZ had a specific activity of 650.13 unitsPHA mg −1 protein. The data reveal that the LIP3, LIP4, and PhaZ enzymes have a wide-range of substrate preferences, including scl-PHA polymers and mcl-PHA polymers, as well as C4 to C10 pNP-alkanoates. These enzymes also showed high activity towards PLA and were also able to hydrolyze synthetic polyester polymers, such as PCL and PES (

Analysis of Degradation Products
We investigated the ability of the LIP3, LIP4, and PhaZ enzymes to degrade PHAs, PLA, and the petrochemical-based polymers, PCL and PES, using Gel Permeation Chromatography (GPC), which enabled quantification of changes in the polymer molecular weight (Mw), mass number (Mn), and molecular mass dispersivity (MP) after incubation with the enzymes. Encouragingly, the polymer samples before and after enzyme treatment showed significant differences. Figures 8-10 show that for all GPC runs, no peak was observed after 11 min of elution. Overlaying the GPC chromatograms of PHBV, PHHx, PHO, PHN, PHD, PLA, PCL, and PES polymers after treatment with LIP3, LIP4, or PhaZ enzymes for 96 h, with chromatograms from untreated polymers, revealed significant changes in Mw, Mn, and MP values (see Figures 8-10 and Supplementary Table S1).
The GPC data also support the observations that the LIP3, LIP4, and PhaZ enzymes were able to hydrolyze ester bonds of a wide variety of polymer types. Table 4-6 compares the percent change in molecular weights of the different polymers tested. These data indicate that molecular weight of PHN reduced by 49.7% after treatment by LIP3 and that LIP3 was able to reduce the molecular weights of PHHx, PHD, PLA, and PCL by similar percentages: 23.4%, 25.9%, 22.5, and 27.9%, respectively ( Table 4). The molecular weight of PCL was reduced by 49.3% by LIP4, which was greater than the reduction in molecular weight of all of the PHA polymers tested and much greater than the reduction in molecular weight of PLA and PES (Table 5). PhaZ was also able to reduce the molecular weight of PCL by 27.6%, which was a far greater reduction in molecular weight caused by PhaZ for all the other polymers tested (Table 6).

Discussion
Analysis of the primary structures of the LIP3, LIP4, and PhaZ enzymes revealed no significant overall sequence homology between them. However, the lipase box consensus sequence motif GX 1 SX 2 G was found, with the X 1 in LIP3 and LIP4 occupied by histidine and asparagine residues, respectively, while PhaZ had valine at the X 1 position. In lipases and esterases, the X 1 residue is usually occupied by a polar residue and X 2 is variable [25]. In contrast, PHA depolymerases have a hydrophobic residue at position X 1 [11,26]. Another conserved sequence region (GMLG-in LIP3, HGGGW-in LIP4 and PFV-in PhaZ) that appears to resemble the oxyanion hole consensus sequence in lipases and depolymerases was observed.
In general, lipases and PHA depolymerases are variable in size, with sequence similarity observable in short structural regions located around the active site residues [11,26]. In this context, Arpigny and Jaeger [27] suggested that bacterial lipases can be organized into eight classes based on their conserved amino acid sequence regions, structural-elements, and biochemical properties. The three-dimensional structures of lipases and PHA depolymerases show that these structures are very similar, despite sequence variability between them. Principally, they characteristically exhibit common motifs in their nuclei, known as α/β-hydrolase folds for the hydrolase and/or esterase families [28]. This common α/β-hydrolase fold consists of an eight-stranded, mostly parallel β-fold flanked by six α-helices, with a catalytic triad of amino acids (Ser/Asp-His-Asp/Glu). Interestingly, LIP3 has seven β-sheets with Ser83-His263-Asp241 in its active site, whereas LIP4 and PhaZ have eight β-sheets and a catalytic triad, Ser180-His307-Asp277 and Ser102-His248-Asp221, respectively.
The LIP3 and PhaZ enzymes were active in a pH range of 4.0 to 10.0 with optimal activity at pH 8 and 45-50 • C. This is consistent with the range reported for nPHAmcl depolymerase from P. putida KT2442 (pH 7.0 to 10.5) [29], as well as the lipase of P. aeruginosa MB 5001, which was reported to have a maximum esterase activity at pH 8.0 and 55 • C [30]. Similar to lipases from the psychrophilic Pseudomonas fluorescens strain AFT 36 [31] and Pseudomonas sp. with pH optima ranging from 6 to 9 [32] and lipases from Pseudomonas sp. and P. fluorescens AFT 36 with thermostabilities below 60 • C [31,33], LIP4 showed maximal activity at pH 6.0 and 50 • C.
Lipase PueB from Pseudomonas chlororaphis [34] and lipase from Pseudomonas sp. [35] showed stability at 100 • C. The thermal stability of LIP3 was relatively higher than that of LIP4 and PhaZ as concluded from the thermal inactivation studies. XtalPred analysis of the amino acid sequence revealed that the secondary structure of LIP3 had a slightly higher percentage of α-helix residues and random coils (88%) than that of LIP4 (82%) and PhaZ (84%). It is known that α-Helices confer conformational stability and random coils impart flexibility to the protein structure, which improves the thermal stability of the enzymes [36,37].
The activity of the LIP3, LIP4, and PhaZ enzymes were neither inhibited nor stimulated by the presence of EDTA and metal ions. These data suggest that metal ions are not required as cofactors for the activities of the three enzymes. The lipases of Trichosporonasteroides strain LP005 [38] and Aspergillus terreus [39] and the PHA depolymerase of P. putida LS46 [36,37] were also not influenced by the presence of metal ions.
The stimulatory effectof Ca 2+ on activity was observed in lipases from Pseudomonas putida 3SK [40], P. aeruginosa EF2 [41], and Pseudomonas sp. 7323 [42]. It has been suggested that calcium ions are primarily involved in the removal of fatty acids formed as insoluble calcium soaps during hydrolysis, resulting in a change in the hydrophobic substratehydrophilic ratio in water [43]. It has also been suggested that the addition of calcium ions alters the orientation of the enzyme on the substrate molecule [44,45].
The inhibition of LIP3, LIP4, and PhaZ by phenylmethanesulphoylflouride (PMSF), a known inhibitor of enzyme activities, was comparable to that of PueB lipase from Pseudomonas chlororaphis, which showed 50% inhibition with 1 mmol L −1 PMSF [34]. The activities of LIP3, LIP4, and PhaZ were also inhibited by ionic and nonionic detergents, sodium dodecyl sulphate (SDS), and Tween 80indicating the presence of a hydrophobic region in the catalytic center and/or a change in the active configuration of the enzyme. Like PMSF, SDS is a known inhibitor of enzyme activities. It is reported to inhibit the lipase of Pseudomonas putida 3SK [40] and the PHA depolymerases of P. fluorescens GK13 [20] and P. putida LS46 [17].
It is known that many lipases exhibit a so-called interfacial activity observable at organic-aqueous interfaces and that many lipases have a mobile subdomain located, referred to as the "lid", above the active site to facilitate this phenomenon [46,47]. The main function of the lid is to protect the active site, and it plays an important role in catalytic activity [48]. In aqueous media, the lid is closed, while in the presence of a hydrophobic layer, it allows access to the binding pocket. The lid of lipase has an amphipathic structure: in the closed state, its hydrophobic side faces the catalytic pocket, while the hydrophilic side faces the solvent [48]. The transition from the closed to the open conformation is accompanied by the exposure of the hydrophobic side to the substrate binding region [49]. Therefore, it is clear that the amphipathic nature of the lid and its specific amino acid sequence are important features for the enzyme activity and specificity of lipases [50]. However, PHA depolymerases and some lipases do not have a lid structure, such as the lipase from Candida antarctica B [15]. Modelling of the three-dimensional structures of LIP3, LIP4, and PhaZ complexed with a 3-hydroxyoctanoate dimer revealed that the core domains of LIP3 and LIP4 are of the α/β hydrolase-type and that their active sites are covered with a closing lid domain, whereas the PhaZdepolymerase lacks the lid domain.
The broad substrate preferences of LIP3, LIP4, and PhaZ have also been reported for the PueB lipase from Pseudomonas chlororaphis, which can also hydrolyze various pNPalkanoates, such as pNP-acetate, pNP-propionate, and pNP-butyrate [46]. Indeed, LIP3 and PhaZ enzymes have the highest activity for pNP-octanoate, but different relative activities are observed for different polyesters. For example, LIP4 showed high relative activity towards pNP-acetate (C2) compared to LIP3 and PhaZ. On the other hand, PhaZ has a comparatively high substrate preference for mcl-PHA polymers. The differences in substrate specificity could be related to the different local microenvironment of the oxyanion pocket as well as to the configuration of the active site of the enzymes, which generate different affinities, activation barriers, and overall energetics of substrate binding.
The GPC data revealed degradation of a wide variety of polymer types by LIP3, LIP4, and PhaZ. A significant reduction in molecular weights of medium chain PHAs was observed with LIP3 (49.7%), while LIP4 and PhaZ exhibited significant degradation of PCL (49.3% and 27.6%) compared to other polymers, which clearly shows its high degradation potential compared to the known lipases/esterases. Kanmani et al. [51] reported a 21% molecular weight decrease of PHAs treated with Bacillus subtilis lipase, while PHA depolymerase from P. putida [17] and other lipases from P. chlororaphis [52] demonstrated 18-40% reduction in molecular weights of polymers after treatment with the enzymes.
PHAs are natural polymers synthesized by bacteria as carbon and energy storage compounds [53]. PHAs exists in two phases, amorphous and crystalline, similar to that observed for the bioceramics with highly variable (supervariate) compositions [54]. Intracellular native PHA polymer granules exhibit an amorphous state, while the PHA chains of extracted granules adopt crystalline phase [55].
The most prominent features of LIP3 and LIP4 that distinguishes them from other lipases described in the literature is their broad substrate specificity ( Table 7). The enzymes LIP3, LIP4, and PhaZ were able to hydrolyze both PCL and PES to some extent. PCL is an aliphatic polyester consisting of repeating ω-hydroxyalkanoate units. Cell-free extracts of the yeast Pseudozyma japonica Y7-09 showed significant ability to degrade PCL [56]. Extracellular lipases of P. aeruginosa have shown the degradation of aromatic-aliphatic polyesters like PCL [1]. PES is a poly(alkene dicarboxylate) polymer structure, and while no studies in the published literature have reported degradation of PES by lipases [57], the intracellular, mcl-PHA depolymerase from P. putida LS46 was shown to have detectable activity towards PES [17].

Bacterial Strains and Plasmids
P. chlororaphis PA23 is a plant growth-promoting bacterium isolated from the rhizosphere of soybean that can suppress the growth of the fungal pathogen Sclerotinia sclerotiorum due to its ability to synthesize phenazine and pyrrolnitrine compounds [2,3]. Escherichia coli DH5a and E. coli BL21 (DE3) were used as host cells for recombinant plasmid construction and protein expression, respectively. Plasmid pET28a(+) (Novagen, Madison, WI, USA) was used as an expression vector for the expression of lip3, lip4, and phaZ. The PHA polymers used in the study were synthesized by Pseudomonas putida LS46 (International Depository Authority of Canada Accession Number 181110-03) [65,66].

DNA Manipulation and Plasmid Construction
The genomic DNA of P. chlororaphis PA23 was isolated from LB bacterial cultures using the Wizard DNA purification kit (Promega). DNA fragments containing genes encoding lipases (lip3 and lip4) andPHA dpolymerase (phaZ) were generated separately by PCR amplification of P. chlororaphis PA23 chromosomal DNA using Phusion DNA polymerase (Thermo Scientific, Waltham, MA, USA),and the primer combinations are listed in Table 9. The nucleotide sequences of these genes were obtained from the whole genome sequence of P. chlororaphis PA23 from NCBI (Accession No. CP008696). Primers were designed to incorporate the restriction endonuclease sites NheI and HindIII at the 5 -ends of the forward and reverse primers, respectively. The signal peptide encoding region of lip4 was not considered in the design of the primers. Table 9. Primers for the construction of recombinant lip3, lip4, and phaZ in the pET28a vector.

Gene
Primers Used
4.6. Qualitative and Quantitative Estimation of Depolymerase/Esterase Activity of LIP3, LIP4, and PhaZ To determine the depolymerase activity of the recombinant enzymes, a homogeneous latex suspension of PHA polymers synthesized by P. putida LS46 from hexanoic (PHHx), octanoic (PHO), nonanoic (PNO), and decanoic acids (PHD) was prepared as described by [29]. Four volumes of a PHA solution in acetone (0.1%, w/v) were added dropwise to one volume of cold water (5-10 • C) with stirring. The acetone was removed with speed vacuum concentrators to obtain a white colloidal suspension. PHA agar plates were prepared using 10 mg PHA polymer suspension and 1.5% (wt/vol) agar in 50 mmol L −1 phosphate buffer (pH 7). The enzyme solution was dropped into 5-mm-diameter wells punched into the PHA agar plates and then incubated at 30 • C for 48 h. The free (clear) zone around the wells indicated depolymerase activity. The fluorescent dye Rhodamine B (0.0005%) was added to the plates to show fluorescent halos around the wells when irradiated with UV light (λ = 350 nm) [77].
The PHA depolymerase activity of the enzyme was determined by measuring the decrease in turbidity of the PHA suspension at OD 650 [62]. The reaction mixture contained 1 mg PHA latex, 50 mmol L −1 phosphate buffer, pH 7, and 0.025 mg protein solution. The turbidity decrease was measured every 24 h for 96 h. One unit of depolymerase activity by turbidimetric assay was determined as the amount of enzyme that can decrease the absorbance (OD 650 ) by 1 absorbance unit per min (i.e.,1OD 650 /min). Alternatively, one unit of depolymerase activity (units PHA ) was the amount of enzyme that hydrolyzed 1 µg of PHA in 1 min. Parallel PHA control samples (without the enzyme) were taken. All experiments were performed in triplicate.
The esterase activity of the enzymes was determined as described by Schirmer et al. [20] using p-nitrophenyl (pNP)-alkanoates with different carbon chain lengths as substrate. The recombinant enzymes from crude cell lysates and/or the purified enzymes (0.025 mg) were used in reactions with 3.9 mL PBS buffer and 100 µL substrate, p-NPalkanoates (1 mmol L −1 ) at optimum temperature for 20 min. The reaction was stopped with 1 mL of 1 M sodium carbonate and the absorbance was recorded at OD 420 . One unit of esterase activity was measured as the amount of enzyme releasing 1 µmol of p-nitrophenol per min under optimal conditions. Total protein was determined by the Bradford assay, using Bovine Serum Albumin (BSA) as the standard protein [78].

Biophysical and Biochemical Properties and Substrate Specificity of LIP3, LIP4 and PhaZ
To determine the effect of pH on the activity of LIP3, LIP4, and PhaZ, the substratep-NP-octanoate (PNPO, 1 mmol L −1 ) was dissolved separately in buffers with different pH values [sodium acetate buffer (50 mmol L −1 , pH 4.0-5.0), sodium phosphate buffer (50 mmol L −1 , pH 6.0-8.0) and glycine-NaOH buffer (50 mmol L −1 , pH 9.0-10.0)] and then used in reaction mixtures with the appropriately diluted enzyme (0.025 mg) solutions prepared in the desired buffers. The effects of temperature on enzyme activity were studied by testing at different temperatures (25-55 • C) for 20 min. The enzymes were exposed to different temperatures (30-60 • C) for a period of 24 h (h) to check their thermal stability. The activation energy of thermal inactivation for the enzymes was calculated from the slope (−E a /R) of Arrhenius plots (k inact = Ae −Ea/RT ), where A represents the Arrhenius constant, E a is the activation energy, R is the gas constant, and T is the absolute temperature.
The effect of cations (chloride/sulfate salts), chelators (EDTA), and ionic and non-ionic detergents on the enzyme activities of LIP3, LIP4, and PhaZ was investigated by adding them to the reaction mixtures and incubating them in 50 mmol L −1 sodium phosphate buffer at the optimal pH and temperature conditions for 20 min. The residual enzyme activities were determined.
Substrate specificity of the enzymes was estimated for PHAs, PHBV, PLA, p-NPalkanoate substrates (pNP-acetate, pNP-butyrate, pNP-octanoate, and pNP-decanoate), and petrochemical based polyester polymers, such as PES and PCL, at a substrate concentration of 1 mg under optimal conditions for enzyme activity. The V max and K m values were calculated using the Lineweaver-Burk linear regression plot. The purified enzymes, LIP3, LIP4, and PhaZ were incubated with pNP-octanoate substrate in concentrations ranging from 0.1 to 3.0 mmol L −1 under optimal assay conditions.

Gel Permeation Chromatography
The molecular weights of PHAs before and after degradation with LIP3, LIP4, and PhaZ were analyzed by gel permeation chromatography (GPC). The reaction mixture with different 10 mg polymer suspension (PHB, PHHx, PHO, PHN, PHD, PLA, PES, and PCL), prepared according to Schirmer et al. [20], was enzymatically hydrolyzed with the recombinant enzymes (2.5 mg) in separate experiments. The polymer substrates (without the enzyme) served as controls. After 96 h of incubation at optimal temperature, the reaction mixtures and substrate controls were oven dried at 60 • C. After cooling, samples were dissolved in chloroform to achieve a final sample concentration of 1.5 mg/mL and filtered with 0.45 mm PTFE. All test and control sample sets were analyzed by gel permeation chromatography (GPC) using a Waters Model 1515 solvent pump with a Waters Refractive Index detector (model 2414) and the Agilent PLgel MIXED-C column (7.5 mm i.d.: 1.5 mL/min). Data were acquired and analyzed using the Breeze 2 software, version (Waters Chromatography, Kent, UK). The procedure was calibrated with Agilent Polystyrene EasiCal PS-1 standards (Agilent Technologies, Mississauga, ON, Canada). The mobile phase (HPLC grade chloroform) was run at a flow rate of 1 mL/min. The column temperature was set at 30 • C and the sample injection volume was set at 20 µL. The method was integrated with the standards to obtain peak-average molecular weight (MP), number-average molecular weight (Mn), weight-average molecular weight (Mw), and polydispersity index (Mw/Mn).

Conclusions
In the present study, the remarkable versatility of the lipases and PHA depolymerase in terms of their potential application to the degradation of various polyesters, including pNPalkanoates, PHAs, PLA, and to some extent the petrochemical-based polyester polymers, PCL and PES, have been demonstrated. Our data suggest that LIP3, LIP4, and PhaZ enzymes differ significantly in their biochemical and biophysical properties, structural folding with the substrate, and absence or presence of the lid domain.

Informed Consent Statement: Not applicable.
Data Availability Statement: All data related to this study are presented in the figures, tables, and Supplementary Materials provided.

Conflicts of Interest:
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.